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BOR - Papers in Press, published online ahead of print October 1, 2003.
Biol Reprod 2003, 10.1095/biolreprod.103.018374
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BIOLOGY OF REPRODUCTION 70, 222–228 (2004)
DOI: 10.1095/biolreprod.103.018374
© 2004 by the Society for the Study of Reproduction, Inc.

Pregnancy

Depression by Relaxin of Neurally Induced Contractile Responses in the Mouse Gastric Fundus1

Maria Caterina Baccari2, Silvia Nistri, Silvia Quattrone, Mario Bigazzi, Tatiana Bani Sacchi, Franco Calamai, and Daniele Bani

Departments of Physiological Sciences3 and Anatomy,4 Histology and Forensic Medicine, University of Florence, Florence, Italy Prosperius Institute,5 Florence, Italy


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The peptide hormone relaxin, which attains high circulating levels during pregnancy, has been shown to depress small-bowel motility through a nitric oxide (NO)-mediated mechanism. In the present study we investigated whether relaxin also influences gastric contractile responses in mice. Female mice in proestrus or estrus were treated for 18 h with relaxin (1 µg s.c.) or vehicle (controls). Mechanical responses of gastric fundal strips were recorded via force-displacement transducers. Evaluation of the expression of nitric oxide synthase (NOS) isoforms was performed by immunohistochemistry and Western blot. In control mice, neurally induced contractile responses elicited by electrical field stimulation (EFS) were reduced in amplitude by addition of relaxin to the organ bath medium. In the presence of the NO synthesis inhibitor L-NNA, relaxin was ineffective. Direct smooth muscle contractile responses were not influenced by relaxin or L-NNA. In strips from relaxin-pretreated mice, the amplitude of neurally induced contractile responses was also reduced in respect to the controls, while that of direct smooth muscle contractions was not. Further addition of relaxin to the bath medium did not influence EFS-induced responses, whereas L-NNA did. An increased expression of NOS I and NOS III was observed in gastric tissues from relaxin-pretreated mice. In conclusion, the peptide hormone relaxin depresses cholinergic contractile responses in the mouse gastric fundus by up-regulating NO biosynthesis at the neural level.

mechanisms of hormone action, nitric oxide, relaxin


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Growing evidence indicates that the peptide hormone relaxin, secreted primarily by the corpus luteum during pregnancy [1, 2], beyond well-demonstrated actions on reproductive tissues [3], exerts its effects on a variety of tissues and organs, thereby affecting several physiological functions. Relaxin, structurally related to insulin and insulin-like growth factor, influences the brain and regulates pituitary hormone secretion [4–6], causes renal vasodilatation by reducing myogenic reactivity [7, 8], increases coronary flow [9], and exerts chronotropic action on the heart [10]. Relaxin also exerts relaxant effects on smooth muscle: in several tissues the hormone appears to act by promoting nitric oxide (NO) biosynthesis [11–13].

NO is synthesized from L-arginine under the catalytic action of NO synthases (NOS) by almost all mammalian cells [14, 15]. Three distinct enzyme isoforms have been identified. Two of them are constitutive and are expressed by endothelial cells (eNOS, type III) or by neurons of the brain and the enteric nervous system (nNOS, type I). The third isoform is a high-yield inducible enzyme (iNOS, type II) that can be either expressed constitutively or induced by a variety of stimuli, including proinflammatory mediators and hormones.

NO released by nonadrenergic, noncholinergic (NANC) nerves is considered an important inhibitory neurotransmitter that can depress cholinergic contractions in the gut [16–19] and also plays a pilot role in relaxation of gastric fundus [20–24] and accommodation of the stomach to food and fluid intake in both animals and humans [25–27]. In vitro studies have shown that NOS activity is increased in the gastrointestinal tract during pregnancy [28, 29]. Recently, it has been reported that relaxin possesses specific binding sites in smooth muscle cells of the small intestine in pregnant pigs [30] and that this hormone is able to depress the myogenic contractile activity of the mouse ileum through an NO-mediated mechanism [13].

The present study was designed to extend the knowledge on the effects of relaxin on the gastrointestinal apparatus by investigating its actions on neurally induced contractile responses in the mouse gastric fundus.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals and Treatments

Virgin albino female mice of Swiss strain, 8–12 wk old and weighing about 30 g (Morini, Reggio Emilia, Italy) were employed. The mice were fed standard laboratory chow and water and were housed under a 12L:12D photoperiod and controlled temperature (21 ± 1°C). The experimental protocol was designed in compliance with the Principles of Laboratory Animal Care (NIH Publications 86-23, revised 1985) and the recommendations of the European Economic Community (86/609/CEE). After 1-wk acclimatization, the mice underwent assessment of the phase of the estrous cycle by light microscopic examination of vaginal smears stained with Papanicolaou, according to Austin and Rowlands [31]. Only mice in proestrus or estrus, that is, the estrogen-dominated phases, entered the experiments. The reason for this choice is that, in several relaxin target organs and tissues, estrogens are required to induce relaxin responsiveness [32, 33]. The mice were randomly distributed in two groups, 18 animals each. The mice of the first group received a single subcutaneous injection of 1 µg of highly purified porcine relaxin (2500–3000 U/mg), prepared according to Sherwood and O'Byrne [34]. As previously done [12, 13], the hormone was dissolved in 0.2 ml of benzopurpurin (Fluka AG, Buchs, Switzerland) 1% in phosphate-buffered saline (PBS), a repository vehicle that allows a slow release of the hormone over 24 h. The chosen dose, vehicle, and route of administration of relaxin were similar to those used in previous studies in mice and proved effective in inducing clear-cut responses in target organs in terms of cell growth [33] or NOS induction [12, 13]. The mice of the second group received the vehicle alone. They are referred to as control mice. Eighteen hours later, the mice were killed by cervical dislocation, and the stomach was rapidly dissected free from the abdomen. An 18-h exposure to relaxin of mice in proestrus or estrus was deemed adequate to obtain a prolonged stimulation [13]. Taking into account the duration of the different phases of the estrous cycle in mice [35], this exposure time also allows the animals not to enter diestrus, the progesterone-dominated phase.

Functional Studies

The stomach was placed on a Petri dish containing warm (37°C) Krebs-Henseleit solution of the following composition: 118 mM NaCl, 4.7 mM KCl, 1.2 mM MgSO4, KH2PO4, 25 mM NaHCO3, 2.5 mM CaCl2, and 10 mM glucose, gassed with 95% O2: 5% CO2 (pH 7.4). Two full-thickness tissue strips (2 x 10 mm) were cut from each fundus region in the direction of the longitudinal muscle. One end of each strip was tied to a platinum rod, while the other was connected to a force displacement transducer (Grass model FT03, Quincy, MA) by a silk thread for continuous recording of isometric tension. The transducer was coupled to a polygraph (Sanborn model 7700, Waltham, MA). The fundal strips were mounted in 5-ml double-jacketed organ baths containing Krebs-Henseleit solution gassed with 95% O2: 5% CO2 mixture. Prewarmed water (37°C) was circulated through the outer jacket of the tissue bath via a constant temperature circulator pump. The temperature of the Krebs-Henseleit solution in the organ bath was maintained within ±0.5°C. As previously reported [24, 36], tissues were allowed to equilibrate for 1 h under an initial load of 0.8 g. During this period, repeated and prolonged washes of the preparations with Krebs-Henseleit solution were done to avoid accumulation of metabolites in the organ baths.

Electrical field stimulation (EFS) was applied via two platinum wire rings (2 mm diameter, 5 mm apart) through which the fundal strip was threaded. Electrical impulses (rectangular waves, 80 V, 4–16 Hz, 0.5 or 5 msec, for 15 sec) were provided by a Grass model S8 stimulator (Grass, Quincy, MA).

Drugs

The following drugs were used: the muscarinic receptors antagonist atropine sulfate (1 x 10-6 M), the cholinergic agonist acetyl-ß-methylcholine chloride (methacholine, 1 x 10-8–1 x 10-3 M), the adrenergic blocker guanethidine monosulfate (1 x 10-6 M), the neural blocker tetrodotoxin (TTX, 1 x 10-6 M), the NOS inhibitor NG-nitro-L-arginine (L-NNA, 2 x 10-4 M), and relaxin (3 x 10-8 M). All drugs were obtained from Sigma (St. Louis, MO), except for relaxin (pure porcine relaxin, 2500–3000 U/mg), which was generously provided by Dr. O.D. Sherwood, University of Illinois (Urbana, IL). Solutions were prepared on the day of the experiment, except for TTX, for which a stock solution was kept stored at -20°C. Drug concentrations are given as final bath concentrations. The concentrations of L-NNA and relaxin used were in the range of those previously shown to be effective in mice to inhibit NO-mediated gastric relaxation [24, 36] and ileal and myometrial contractile activity [12, 13], respectively. Drugs were dissolved in Krebs-Henseleit solution and were added in volumes not exceeding 0.5% of the bath volume.

Neurally evoked and directly induced muscular responses by EFS have been elicited in fundal strips from control mice both in the absence and 20–30 min after the addition of guanethidine, TTX, or atropine to the bath medium. The effects of RLX or L-NNA on neurally induced contractions to EFS (4–16 Hz, 0.5 msec) were first investigated separately on different strips, starting 10 min after their addition to the bath medium, then in combination to study the reciprocal influence of the two substances on the contractile responses themselves. In these latter experimental conditions, L-NNA was added to the bath medium 20 min following RLX, or, in a different set of strips, RLX was added to the bath medium 10–15 min following L-NNA. Direct smooth muscle responses, besides EFS (4–16 Hz, 5 msec), were also evoked by adding methacholine (2 x 10-6 M) to the bath medium either in the absence or 20 min following addition of TTX. The interval between two subsequent applications of methacoline was no less than 30 min, during which repeated and prolonged washes of the preparations with Krebs-Henseleit solution were performed. Methacholine (1 x 10-8–1 x 10-3 M) concentration-response curve was also obtained cumulatively with a stepwise increase in concentration once a stable plateau was reached with the previously added concentration. The effects of RLX or L-NNA on direct smooth muscle responses were investigated in the presence of TTX or atropine when EFS was performed and in the presence of TTX only when methacholine was employed.

In order to assess the viability of the preparations, contractile responses to 2 x 10-6 M methacholine were performed and compared at the beginning and at the end of each experiment, except in the strips that had been treated with atropine.

The previously described series of experiments were also carried out on fundal strips from relaxin-pretreated mice.

Immunohistochemistry for NOS Isoform Localization

Stomach fragments from the control and the RLX-treated mice were fixed by immersion in paraformaldehyde 4% in PBS (pH 7.4) for 2 h, cryoprotected by incubation in sucrose 20% in PBS for 12 h, and washed in PBS and quickly frozen at -80° C in cryostat embedding medium (Bio-Optica, Milan, Italy). Cryostat sections, 6 µm thick, were cut from each tissue fragment and immunostained with rabbit polyclonal antibodies against NOS III (Alexis, Läufelingen, Switzerland), NOS II (Alexis), or NOS I (Calbiochem, San Diego, CA). The antisera were applied to sections at working dilutions of 1:50 (anti-NOS I and III) or 1:100 (anti-NOS II) in PBS overnight at 4°C. Negative controls were carried out by omitting the primary antisera or by preabsorbing the specific anti-NOS antibodies with corresponding blocking peptides (Calbiochem) following the protocol indicated by the manufacturer. The immune reaction was revealed by Alexa-488nm-labeled goat anti-rabbit antibodies (Molecular Probes; diluted 1:200 in PBS). To achieve a precise intracellular localization of NOS isoforms, the immunostained sections were examined with a Bio-Rad 1024 ES confocal laser scanning microscope (Bio-Rad, Hempstead Herts, UK) with laser beam excitation at 488-nm wave length.

Evaluation of NOS Expression by Western Blot

Stomach fragments from the control and the RLX-treated mice were quickly minced and homogenized with a tissue homogenizer (Ing. Terzano, Milan, Italy) in 500 µl cold lysis buffer of the following composition: 20 mM Tris/HCl (pH 7.4), 10 mM NaCl, 1.5 mM MgCl2, 5 mM EGTA, 2 mM Na2EDTA, 1 mM dithiotreitol (DTT), 1 mM phenylmethylsulfonyl fluoride (PMSF), 1% Triton X-100, 20 µg/ml leupeptin, 1 µg/ml pepstatin, 500 µg/ml pefabloc, and 2.5 µg/ml aprotinin. On centrifugation at 17 000 x g at 4°C, the supernatants were collected, and the total protein content was measured spectrophotometrically using Bradford reagent (Sigma). Samples of the supernatants, each containing 80 µg of proteins, and appropriate molecular weight markers (Bio-Rad, Hercules, CA) were electrophoresed by SDS-PAGE (200 V, 1 h) using a denaturating 7.6% polyacrylamide gel and blotted onto nitrocellulose membranes (Amersham Pharmacia Biotech Italy, Cologno Monzese, Italy; 150 V, 1 h). After thorough washings in PBS containing 0.1% Tween (T-PBS), the membranes were blocked with 20 ml T-PBS containing 5% bovine serum albumin (BSA; Sigma) at room temperature for 1 h and incubated overnight at 4°C with stirring in the same rabbit polyclonal antisera used for immunohistochemistry, which were diluted 1:50 000 in T-PBS containing 1% BSA. The immune reaction product was revealed by incubating membranes in peroxidase-labeled goat anti-rabbit antibodies (Vector, Burlingame, CA), diluted 1:10 000 in T-PBS containing 1% BSA for 1 h at room temperature with stirring. After a brief wash the membranes were incubated with the chemiluminescent substrate ECL (Amersham) and then exposed to high-sensitivity photographic film (Biomax ML, Kodak, Rochester, NY) for varying exposure times in order to identify the correct film exposure that did not reach maximum for any of the signals (usually 1 min). After stripping the bound anti-NOS immune complexes with a solution containing 62.5 mM Tris-HCl, 100 mM ß-mercaptoethanol, and 2% SDS (30 min at 50°C), the membranes were immunostained with rabbit polyclonal anti-pan-actin antibodies (Zymed, San Francisco, CA; diluted 1:20 000 in T-PBS containing 1% BSA) using the same procedure as for NOS immunostaining. Actin was assumed as control invariant protein whose expression should not vary on relaxin treatment. For each NOS isoform, quantitative evaluation of the bands appearing on the photographic film was performed by computer-assisted densitometry. The bands, each corresponding to an individual mouse, were digitized with a flatbed scanner, and their optical density was measured using the Scion Image Beta 4.0.2 image analysis program (Scion Corp., Frederick, MD).

Data Analysis and Statistical Test

Amplitude of contractile responses to EFS is expressed as percentage of the muscular contraction evoked by 2 x 10-6 M methacholine, assumed as 100%. Amplitude of contractile responses to methacholine was measured 30 sec after a stable plateau phase was reached. Percentage data were arcsine square root transformed before being statistically analyzed. All values are expressed as mean ± SEM. The number of preparations is designated by n in the Results section.

Statistical analysis was performed by means of Student t-test to compare two experimental groups or of one-way ANOVA when more than two groups were compared. When ANOVA indicated that significant differences existed, multiple comparisons between groups were carried out by Newman-Keuls posttest. For all statistical tests, values were considered significantly different with P values equal or less than 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Contractile Responses of Strips from Control Mice

At basal tension, no spontaneous motor activity was present, while EFS (4–16 Hz, 0.5 msec) elicited contractile responses (n = 12) whose amplitude increased by increasing the stimulation frequency (Fig. 1). The EFS-evoked contractile responses were abolished by TTX (1 x 10-6 M) (P < 0.05) or atropine (1 x 10-6 M) (P < 0.05) and were not influenced by guanethidine (1 x 10-6 M) (P > 0.05), thus indicating that they were neurally induced and cholinergic in nature.



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  		FIG. 1. Contractile responses elicited by EFS in the gastric fundus from control mice. A) Typical tracings showing contractile responses to EFS. Compared to the control responses (left record), addition of relaxin (3 x 10-8 M, center record) causes a reduction in amplitude of the EFS-induced contractions in the whole range of stimulation frequency employed. The NO synthesis inhibitor L-NNA (2 x 10-4 M), added to the bath medium 20 min after relaxin (right record), markedly increases the amplitude of the EFS-induced contractile responses. The three records are obtained from the same strip. B) Graphical representation of the mean amplitude of EFS-induced contractile responses at different stimulation frequencies. The amplitude of the contractile responses is expressed as percentages of the contraction induced by methacholine (2 x 10-6 M), assumed as 100%. Relaxin, 20 min after its addition to the bath fluid, decreases the mean amplitude of the contractile responses in the whole range of stimulation frequency employed. Subsequent addition of L-NNA to the bath medium in the presence of relaxin causes, after 10 min, a significant increase in the mean amplitude of the EFS-induced contractile responses in respect to the controls. All values are means ± SEM of six preparations. *P < 0.05 versus the controls (one-way ANOVA and Newman-Keuls posttest)

Addition of relaxin (3 x 10-8 M) to the bath medium (n = 6) did not affect the basal tension but caused a clear-cut reduction in amplitude of the EFS-induced contractile responses in the whole range of stimulation frequency employed (Fig. 1). The effects of relaxin, already appreciable 10 min after its addition to the bath medium, were fully evident after 25–30 min and persisted up to 1 h (longer time not observed).

Addition of the NOS inhibitor L-NNA (2 x 10-4 M) to the bath medium (n = 6) caused an increase in amplitude of the EFS-induced contractions, in the whole range of stimulation frequency employed, which was already present 10 min after its addition to the bath medium and persisted up to 1 h (longer time not observed). The effect of L-NNA was also observed in the presence of relaxin (Fig. 1). On the other hand, relaxin (3 x 10-8 M), added to the bath medium 10 min after L-NNA, when the effects of the NOS inhibitor were already manifested, was no more able to reduce the amplitude of the EFS-induced contractile responses (P > 0.05) (data not shown).

EFS, applied at 5-msec pulse duration (n = 8), evoked contractile responses that persisted in the presence of TTX (1 x 10-6 M) (P > 0.05) and/or atropine (1 x 10-6 M) (P > 0.05), thus indicating a direct activation of the smooth muscle [37]. Relaxin (3 x 10-8 M) or L-NNA (2 x 10-4 M) were unable to influence these directly evoked muscular contractions (data not shown).

Addition of the muscarinic receptors agonist methacholine (2 x 10-6 M) to the bath medium caused a sustained contracture after 10–15 sec of contact time, which suddenly reached a plateau (0.76 ± 0.2 g) that persisted until washout (2 min, longer time not observed). Relaxin (3 x 10-8 M) (n = 4) did not influence (P > 0.05) the response to methacholine (Fig. 2). The amplitude of the response to methacholine was not affected by TTX (1 x 10-6 M) (P > 0.05) or L-NNA (2 x 10-4 M) (P > 0.05) (data not shown).



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  		FIG. 2. Mean amplitude of direct smooth muscle contractile responses elicited by methacholine. Mean amplitude of methacholine (1 x 10-8–1 x 10-3 M)-induced concentration-dependent contractions in strips from both control mice, before (open circles) and 20 min after the addition of relaxin to the bath medium (black circles), and RLX-pretreated mice (down triangles). Contractions are expressed as grams. All values are means ± SEM of eight preparations. P > 0.05 for all comparisons versus the controls (one-way ANOVA)

The effects of relaxin (3 x 10-8 M) on the concentration-response curve to methacholine (n = 8) were also evaluated. Methacholine (1 x 10-8–1 x 10-3 M) induced concentration-dependent contractions whose amplitude was not affected by relaxin (Fig. 2).

Contractile Responses of Strips from Relaxin-Pretreated Mice

At basal tension, contractile responses evoked by EFS (4–16 Hz, 0.5 msec) in the strips from relaxin-pretreated mice (n = 12) were greatly reduced in amplitude (n = 10) or even absent (n = 2) as compared with the control animals (P < 0.05), in the whole range of stimulation frequency employed (Fig. 3A; compare with Fig. 1A).



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  		FIG. 3. Contractile responses elicited by EFS in the gastric fundus from relaxin-pretreated mice. A) Typical tracings showing contractile responses to EFS. Contractile responses (left record) appear greatly reduced in amplitude compared to those obtained in strips from control mice (Fig. 1A, left record). Relaxin (3 x 10-8 M, center record) does not influence the amplitude of the contractile responses. The NO synthesis inhibitor L-NNA (2 x 10-4 M), added to the bath medium 20 min after relaxin (right record), potentiates the amplitude of the EFS-induced contractile responses. The three records are obtained from the same strip. B) Graphical representation of the mean amplitude of EFS-induced contractile responses at different stimulation frequencies. The amplitude of the contractile responses is expressed as percentages of the contraction induced by methacholine (2 x 10-6 M), assumed as 100%. Relaxin, 20 min after its addition to the bath fluid, does not influence the mean amplitude of the EFS-induced contractile responses in the whole range of stimulation frequency employed. Subsequent addition of L-NNA to the bath medium in the presence of relaxin still potentiates, after 10 min, the amplitude of EFS-induced contractions. All values are means ± SEM of six preparations. *P < 0.05 versus the controls (one-way ANOVA and Newman-Keuls posttest)

In those strips in which excitatory responses were present, addition of relaxin (3 x 10-8 M) to the bath fluid (n = 8) did not further reduce the amplitude of the EFS-induced contractile responses (Fig. 3).

Addition of L-NNA (2 x 10-4 M) to the bath medium (n = 6) potentiated the amplitude of the EFS-induced contractile responses (P < 0.05) even in the presence of relaxin (P < 0.05) (Fig. 3).

Responses to methacholine, added to the bath medium either as a single dose (2 x 10-6 M) or as increasing concentrations (1 x 10-8–1 x 10-3 M), were not different in amplitude (P > 0.05) as compared with the control mice, thus indicating that, in preparations from relaxin-pretreated animals, contractility was not impaired (Fig. 2). Addition of further relaxin (3 x 10-8 M) to the bath medium did not influence responses to methacholine.

Immunocytochemical Localization of NOS Isoforms

Confocal laser scanning microscopy allowed us to identify the different NOS isoforms in the gastric wall (Fig. 4). Fair NOS I immunoreactivity was almost exclusively detected in structures featuring neurons and nerve fibers of the intramural neural plexuses. NOS II was expressed by squamous cells of the fundal epithelium, by exocrine cells of the gastric glands, and, to a lesser extent, by smooth muscle cells of the muscle coat. NOS III was recognized in smooth muscle cells of the gastric muscle coat and in endothelial cells of blood vessels. With the method used, no clear-cut changes in NOS distribution and staining intensity were observed between the control and the relaxin-treated mice apart from an apparent increase in the amount of NOS I neurons and nerve fibers.



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  		FIG. 4. Representative micrographs of whole-thickness gastric specimens from control and relaxin-pretreated mice immunostained to reveal NOS I, NOS II, and NOS III. NOS I is localized chiefly in neurons and nerve fibers of the intramural neural plexuses: specimens from control mice appear less intensely labeled than those from relaxin-treated mice. NOS II labels mainly smooth muscle cells of the circular (CML) and longitudinal (LML) muscle layers, squamous cells of the surface epithelium (not shown), and gland cells (both panels are from specimens of relaxin-treated mice). NOS III is localized in smooth muscle cells of the circular (CML) and longitudinal (LML) muscle layers and in blood vessel endothelium, indicated by arrows (both panels are from specimens of relaxin-treated mice). Confocal microscopy. Bar = 50 µm

Quantitative Expression of NOS Isoforms

By Western blot analysis (Fig. 5), the amounts of NOS I and NOS III proteins expressed by the stomach of the relaxin-pretreated mice were significantly higher than in the control mice. Conversely, there were no significant differences in the amount of NOS II between the two experimental groups. Expression of actin as control household protein appeared unchanged on relaxin treatment.



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  		FIG. 5. Western blot analysis of the expression of NOS isoforms by the stomach of control and relaxin-treated mice. Actin expression is also shown as a control invariant protein for loading equivalency. Upper panel: representative bands from a typical experiment. Lower panel: histogram showing densitometric analysis of the bands. Open columns: control mice. Dark columns: relaxin-treated mice. Relaxin significantly increases the expression of NOS I and NOS III but not NOS II (each column, n = 6 preparations). *P < 0.001 versus the controls (Student t-test)


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The present study shows that the peptide hormone relaxin, which has been previously reported to markedly depress spontaneous motility of the mouse ileum [13], also reduces neurally induced contractile responses of the gastric fundus.

At basal tension, longitudinal strips of gastric fundus appeared mechanically quiescent and did not show spontaneous contractions, as reported in gastric whole-organ in vitro preparations in mice [38]. The lack of spontaneous motor activity in the fundal strips reflects the functional role of this gastric region and implies that activation of muscle contractions in the fundus is dependent on external control, neural, and/or hormonal [39, 40]. Contractile responses can be easily obtained in fundal gastric strips by means of EFS. These motor responses resembled those observed in other animal species [41, 42] and appeared to be caused by activation of excitatory cholinergic nerves. In fact, the neural blocker TTX and/or the muscarinic receptor antagonist atropine always blocked the EFS-evoked motor responses. The reduction in amplitude of the neurally induced contractile responses by relaxin and the lack of effects of the hormone on direct smooth muscle responses, observed in strips from either control or relaxin-pretreated mice, suggest that in gastric fundus the depressant effects of relaxin on EFS-induced contractions are exerted at the nervous level in the nerve-muscle pathway.

EFS stimulates both excitatory and inhibitory nerves, and an inhibitory modulation by NO on cholinergic nerves has been widely demonstrated in the gastrointestinal tract [16–19] and elsewhere [43–45]. In the present experiments, the NO synthesis inhibitor L-NNA enhanced the amplitude of the EFS-induced contractions without affecting direct smooth muscle contractile responses. L-NNA also abolished the depressant effects of relaxin on the EFS-induced contractions, thus suggesting that this NO synthesis inhibitor was able to remove a nitrergic inhibitory influence, stimulated by relaxin itself, on neurally induced contractions. Furthermore, in the presence of L-NNA, relaxin was no longer effective in depressing the neurally induced cholinergic contractions. Therefore, it is conceivable that the depressant effects of relaxin may be exerted through NO biosynthesis and release. The current findings are at variance with those in the ileum of relaxin-pretreated mice, in which we observed an overexpression of NOS II, but not NOS I, that accounted for an intrinsic muscular NO pathway involved in the depressant effects of relaxin [13]. In the current study on the stomach of relaxin-pretreated mice, neuronal NOS I, but not NOS II, was overexpressed in respect to the controls. Taken together, the present data indicate that relaxin acts on the peripheral nervous control of the stomach. These findings fit well with previous studies in the rat brain, in which relaxin was found to bind to and influence the activity of neurons [6], especially in areas rich in nitrergic neurons [46]. All these data offer strong support to the concept that, at the nervous level, relaxin could be a modulator of NO activity. Several substances may stimulate neurogenic NO formation: recently, it has been suggested that nicotinic receptors might mediate NOS expression in the gastric myenteric plexus [47] and that neuronal muscarinic receptors too might be involved in NO formation and release [48, 49]. Indeed, the exact mechanism of action of relaxin on the nervous control of the stomach remains to be elucidated. Nonetheless, based on our previous studies and the current findings, it appears that, in the gastrointestinal tract, relaxin, by up-regulating NO biosynthesis, can depress either myogenic contractions—as occurs in the ileum [13]—or neurally induced contractions, as it does in the stomach. Of note, in pregnant rats, a reduction of gastric motility strictly related to an increased NO release from NANC nerves was observed, whereas such a mechanism is not operating in the ileum [29].

In conclusion, it could be hypothesized that relaxin plays a role in the adjustments of gastrointestinal tract motility occurring in pregnancy: through the depression of contractile responses in the gut, relaxin might increase transit time of ingesta, thereby favoring digestion and absorption of nutrients. In this view, a prolonged transit time caused by relaxin may contribute, together with other hormones that are also increased during gestation, to the altered gastrointestinal motility frequently observed in pregnant state [50].


   ACKNOWLEDGMENTS
 
We thank Prof. B. Piochi for his helpful advice regarding the statistical analyses.


   FOOTNOTES
 
1 This study was supported by funds from the University of Florence, Florence, Italy. The financial support of Telethon-Italy (grant GGP02152) is also gratefully acknowledged. Back

2 Correspondence: Maria Caterina Baccari, Department of Physiological Sciences, University of Florence, Viale G.B. Morgagni 63, I-50134 Florence, Italy. FAX: 39 055 437 9506; mcaterina.baccari{at}unifi.it Back

Received: 17 April 2003.

First decision: 12 May 2003.

Accepted: 12 September 2003.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. O'Byrne EM, Steinetz BG. Radioimmunoassay of relaxin in sera of various species using an antiserum to porcine relaxin. Proc Soc Exp Biol Med 1976 152:272-276[CrossRef][Medline]
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